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Endocrinology Vol. 148, No. 8 3750-3757
Copyright © 2007 by The Endocrine Society

Developmental Control of Plasma Leptin and Adipose Leptin Messenger Ribonucleic Acid in the Ovine Fetus during Late Gestation: Role of Glucocorticoids and Thyroid Hormones

Deirdre M. O’Connor, Dominique Blache, Nigel Hoggard, Emily Brookes, F. B. Peter Wooding, Abigail L. Fowden and Alison J. Forhead

Department of Physiology, Development and Neuroscience (D.M.O., E.B., F.B.P.W., A.L.F., A.J.F.), University of Cambridge, Cambridge CB2 3EG, United Kingdom; School of Animal Biology (D.B.), Faculty of Natural and Agricultural Sciences, University of Western Australia, Perth, Western Australia 6009, Australia; and Obesity and Metabolic Health Group (N.H.), The Rowett Research Institute, Aberdeen AB21 9SB, United Kingdom

Address all correspondence and requests for reprints to: Dr. Alison J. Forhead, Department of Physiology, Development, and Neuroscience, University of Cambridge, Downing Street, Cambridge CB2 3EG, United Kingdom. E-mail: ajf1005{at}cam.ac.uk.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
In developed countries, the increasing incidence of obesity is a serious health problem. Leptin exposure in the perinatal period affects long-term regulation of appetite and energy expenditure, but control of leptin production in utero is unclear. This study investigated perirenal adipose tissue (PAT) and placental leptin expression in ovine fetuses during late gestation and after manipulation of plasma glucocorticoid and thyroid hormone concentrations. Between 130 and 144 d of gestation (term at 145 ± 2 d), plasma leptin and PAT leptin mRNA levels increased in association with increments in plasma cortisol and T3. Fetal adrenalectomy prevented these developmental changes, and exposure of intact 130 d fetuses to glucocorticoids, by cortisol infusion or maternal dexamethasone treatment, caused premature elevations in plasma leptin and PAT leptin gene expression. Fetal thyroidectomy increased plasma leptin and PAT leptin mRNA abundance, whereas intravenous T3 infusion to intact 130 d fetuses had no effect on circulating or PAT leptin. Leptin mRNA expression was low in the ovine placenta. Therefore, in the sheep fetus, PAT appears to be a primary source of leptin in the circulation, and leptin gene expression is regulated by both glucocorticoids and thyroid hormones. Developmental changes in circulating and PAT leptin may mediate the maturational effects of cortisol in utero and have long-term consequences for appetite regulation and the development of obesity.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE ESCALATING INCIDENCE of child and adult obesity in developed countries is a serious health issue and has highlighted the importance of determining the factors that control appetite and energy balance. Some of these factors may act in critical windows of early life. For instance, endocrine signals, such as leptin, in the perinatal period are known to influence the development of neural circuits involved in appetite regulation (1, 2). In rodents, hypothalamic networks concerned with feeding behavior are established in the first 2 wk of life and are dependent on normal exposure to leptin over this period (3). Leptin-deficient ob/ob mice show abnormal development of neural projections from the arcuate nucleus, which can be corrected by leptin treatment in neonatal but not adult life (3). Because the rodent brain develops later than the human and ovine brain, leptin-sensitive formation of the hypothalamic appetite-regulatory network may occur before birth in more precocial animal species (1, 2).

Leptin is detected in the fetal circulation from midgestation in many animal species, including man and sheep (4, 5), and leptin receptors are expressed in a variety of fetal and placental tissues (6, 7). Developmental control of leptin before birth, however, is poorly understood. In human fetuses, plasma leptin concentration rises in late gestation and directly correlates with body weight and ponderal index at delivery (4, 8, 9). Furthermore, in fetal sheep, leptin mRNA abundance in perirenal adipose tissue (PAT) also increases with gestational age, such that adipose tissue in the fetus appears to be the primary source of circulating leptin in utero (10, 11). However, in some animal species, such as man and rats, the placenta is a potential site of leptin synthesis and secretion that may contribute to concentrations in both the fetal and maternal circulations (12).

In the sheep fetus toward term, the rise in plasma leptin has been shown to depend on the coincident rise in plasma cortisol concentration (13). Fetal adrenalectomy, which abolishes the prepartum cortisol surge, prevents the normal rise in circulating leptin in utero, and exogenous infusions of cortisol or dexamethasone have been shown previously to increase plasma leptin concentration in the sheep fetus (13). The prepartum cortisol surge has many maturational effects on the fetus, some of which are mediated by the thyroid hormones (14, 15). Close to term, glucocorticoids induce tissue-specific changes in deiodinase enzyme activity that stimulate an increase in plasma T3 concentration (16). However, the extent to which glucocorticoid-induced changes in plasma leptin are mediated by the thyroid hormones is unknown.

Therefore, the aims of this study were 1) to determine whether developmental and glucocorticoid-induced changes in circulating leptin in utero are attributable to changes in adipose leptin mRNA abundance and 2) to establish the role of thyroid hormones in the control of circulating leptin and leptin mRNA levels in fetal adipose tissue. The study tested the hypothesis that developmental and glucocorticoid-induced changes in plasma leptin before birth are caused by changes in adipose leptin gene expression that, in turn, may be mediated by thyroid hormones. Glucocorticoid and thyroid hormone concentrations in the sheep fetus were manipulated by endocrine gland removal and exogenous hormone treatment.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals
Forty-three Welsh Mountain sheep fetuses of known gestational age were used in this study; 30 were singletons, and 13 were twin fetuses. In total, there were 26 male and 17 female fetuses. The ewes were housed in individual pens and were maintained on 200 g/kg concentrates with free access to hay, water, and a salt-lick block. Food, but not water, was withheld for 18–24 h before surgery. All surgical and experimental procedures were in accordance with the United Kingdom Animals (Scientific Procedures) Act 1986 and were approved by the local animal ethics committee.

Surgical procedures
All surgical operations were performed under halothane anesthesia (1.5% in O2-N2O) with positive pressure ventilation. Five fetuses were thyroidectomized (TX) between 105 and 110 d of gestation, and an additional five fetuses were adrenalectomized (AX) between 116 and 120 d of gestation (term at 145 ± 2 d) using surgical techniques described previously (17, 18). For two of the AX and one of the TX fetuses, the unoperated twin acted as a control intact fetus.

In 15 of the remaining intact fetuses, intravascular catheters were inserted into the femoral artery and vein between 115 and 118 d of gestation (19). All catheters were exteriorized through the flank of the ewe and secured in a plastic bag sutured to the skin. The catheters were flushed daily with heparinized saline solution (100 IU/ml heparin, 0.9% saline wt/vol) from the day after surgery. At surgery, all fetuses were administered 100 mg ampicillin iv (Penbritin; Beecham Animal Health, Brentford, UK) and 2 mg gentamycin iv (Frangen-100; Biovet, Mullingar, Ireland). The ewes were given antibiotics im (procaine penicillin, Depocillin; Mycofarm, Cambridge, UK) on the day of surgery and for 3 d thereafter.

Experimental procedures
Effect of adrenalectomy and thyroidectomy.
The AX (n = 5), TX (n = 5), and untreated intact fetuses (n = 6) were delivered by cesarean section under general anesthesia (20 mg/kg sodium pentobarbitone iv) for collection of tissue and blood samples at 141–146 d of gestation (144 ± 0.3 d); two untreated fetuses were delivered at 129–130 d of gestation. At delivery, 10 ml blood samples were taken by venepuncture of the umbilical artery, and a number of tissues were collected from the fetuses after the administration of a lethal dose of barbiturate (200 mg/kg sodium pentobarbitone). Samples of fetal PAT and placenta were either immediately fixed in 4% paraformaldehyde or frozen in liquid nitrogen and stored at –80 C until analysis. At delivery, there was no evidence of adrenal or thyroid gland remnants in any of the AX or TX fetuses, respectively.

Effect of exogenous cortisol or T3 infusion.
Fifteen chronically catheterized intact fetuses were infused iv with either saline (0.9% NaCl, 3 ml/d; n = 5), cortisol (2–3 mg/kg/d; n = 5; Efcortelan; Glaxo, Ware, UK) or T3 (8–12 µg/kg/d; n = 5; Sigma, Poole, UK) for 5 d before delivery for tissue collection at 129–131 d (130 ± 0.2 d). The doses of cortisol and T3 infused were calculated to mimic the plasma concentrations normally observed in the immediate prepartum period (20). Daily arterial blood samples (4 ml) were taken immediately before and throughout the infusion. At delivery, blood and tissue samples were collected as described above.

Effect of maternal dexamethasone treatment.
From 125 d of gestation, 10 unoperated ewes were injected twice im with either saline (0.9% NaCl; n = 5) or dexamethasone (two times at 12 mg in 2 ml 0.9% NaCl; n = 5) at 24 h intervals. Mean ± SEM body weight was 56 ± 4 kg for the saline-treated ewes and 55 ± 3 kg for the ewes injected with dexamethasone. All of the fetuses were delivered by cesarean section under general anesthesia (20 mg/kg sodium pentobarbitone iv) at 10 h after the second injection. Before anesthesia, a 10 ml blood sample was obtained from the ewe by jugular venepuncture. At delivery, a 10 ml blood sample was taken by venepuncture of the umbilical artery, and a number of tissues, including PAT and placenta, were collected from the ewe and fetus after the administration of a lethal dose of barbiturate (200 mg/kg sodium pentobarbitone). All tissue samples were immediately frozen in liquid nitrogen and stored at –80 C until analysis.

Biochemical and molecular analyses
Plasma hormone concentrations.
All blood samples were immediately placed into EDTA-containing tubes and centrifuged for 5 min at 1000 x g and 4 C. The plasma aliquots were stored at –20 C until analysis. Plasma leptin concentration was measured by RIA using recombinant bovine-ovine leptin standards as described previously (21); the lower limit of detection was 0.09 ng/ml, and the interassay coefficient of variation was 5%. Plasma cortisol concentration was measured by RIA (22) in which the lower limit of detection was 1.0–1.5 ng/ml and the interassay coefficient of variation was 12%. Plasma T3 and T4 concentrations were also measured by RIA using a commercial kit (23). The lower limits of detection were 0.07 ng/ml for T3 and 7.6 ng/ml for T4, and the interassay coefficients of variation were 10% for both assays. All RIAs were validated for use with ovine plasma.

Immunohistochemistry.
The fixed tissue samples were dehydrated in ethanol, transferred to a 50:50 dilution of ethanol and xylene (Histoclear; National Diagnostics, Atlanta, GA) for 1–2 h and then incubated in xylene overnight. Samples were incubated in three changes of paraffin wax at 60 C for 2 h and then blocked out in fresh wax. Sections of 7 µm thickness were cut and mounted on microscope slides. Sections were dewaxed in xylene and transferred to 100% ethanol for 10 min. After washing in distilled water for 20 min, the sections were incubated with PBS/1% BSA/0.05% thiomersal for 5 min and incubated with 5% goat serum/PBS for 1 h to prevent nonspecific antibody binding. The sections were incubated with the primary leptin antibody at 1:1000 (rabbit polyclonal antibody to ovine leptin) or PBS/1% BSA/0.05% thiomersal alone (negative control) overnight at 4 C. Unbound primary antibody was removed by washing with PBS. The sections were incubated at room temperature for 40 min with the secondary antibody (4 nm gold-labeled goat antirabbit; Jackson ImmunoResearch, West Grove, PA) at a dilution of 1:40. All sections were washed in PBS and distilled water and then incubated with silver intensifying solution (IntenSE M silver enhancement kit; GE Healthcare, Little Chalfont, UK) for up to 20 min. The sections were air dried and mounted.

In situ hybridization.
Oligonucleotide probes to ovine leptin were commercially synthesized (Sigma-Genosys, Cambridge, UK). Antisense and sense probes were 5'-ACT GCG TGT GTG AGA TGT CAT TGA TCC TGG TGA CAA TCG TCT TGA TGA GG 3' and 5' CCT CAT CAA GAC GAT TGT CAC CAG GAT CAA TGA CAT CTC ACA CAC GCA GT-3', respectively, and were labeled with digoxigenin (DIG) using DIG oligonucleotide tailing kit (Roche Diagnostics, Welwyn Garden City, UK) according to the instructions of the manufacturer.

Sections of 10 µm thickness were cut from frozen tissue and mounted on poly-lysine-coated microscope slides. Sections were fixed in 4% paraformaldehyde/PBS on ice for 5 min and washed twice for 2.5 min in PBS. The slides were then transferred to 50%, 70% and 95% ethanol for 5 min each. The slides were stored in 95% ethanol at 4 C until needed.

Sections were incubated in prehybridization solution [47% deionized formamide, 2x sodium chloride and sodium citrate (SSC), 10% dextran sulfate, 1x Denhardt’s solution, 50 mM sodium phosphate (pH 7.5), 250 µg/ml salmon sperm DNA, and 500 µg/ml tRNA] for 2 h at 37 C. The sections were then washed in PBS and 70, 95, and 100% ethanol for 3 min each and allowed to air dry. The sections were incubated overnight with hybridization solution (prehybridization solution and labeled probe) at 37 C. Sections were washed at 37 C in 2x SSC, 1x SSC, and 0.5x SSC twice for 15 min each. The slides were washed at room temperature in 100 mM Tris-HCl (pH 7.5), 150 mM NaCl [Tris-buffered saline (TBS)] twice for 10 min each wash. The sections were incubated with blocking solution (TBS, 2% sheep serum, and 0.1% Triton X-100) for 30 min at room temperature and then with the detecting antibody (anti-DIG alkaline phosphate conjugate; Roche Diagnostics) at 1:100 overnight at 4 C. The sections were washed in TBS twice for 10 min each wash and then transferred to 100 mM Tris-HCl (pH 9.5), 100 mM NaCl, 50 mM MgCl2 for an additional 10 min. Nitro blue tetrazolium/5-bromo-4-chloro-3-indolyl phosphate color developing solution [stock solution diluted 1:40 in 100 mM Tris-HCl (pH 9.5), 100 mM NaCl, 50 mM MgCl2] (Roche Diagnostics) was applied to the sections, which were incubated in the dark at room temperature for up to 4 h. The reaction was stopped with the application of 10 mM Tris-HCl (pH 7.5), 1 mM EDTA. Sections were allowed to air dry and were mounted.

Quantitative real-time PCR.
Tissue samples of approximately 100 mg were homogenized on ice in 1 ml Tri reagent (Sigma) using a Polytron homogenizer, and RNA extraction was performed following the method of Chomczynski and Sacchi (24). The RNA samples were resuspended in diethylpyrocarbonate-H2O and were treated with deoxyribonuclease I according to the instructions of the manufacturer (Sigma). The yield of RNA was calculated from the absorbance at 260 nm, and RNA purity was determined by the A260/A280 nm ratio. Samples with a ratio of between 1.8 and 2.0 were considered to be of an acceptable purity. Total RNA (1 µg) was reverse transcribed into cDNA. The reaction was primed with 500 ng oligo-dT (Roche Diagnostics), and the reverse transcriptase used was Superscript II (Invitrogen, Paisley, UK); the reactions were performed according to the instructions of the manufacturer.

Oligonucleotide primers for leptin, cyclophilin, and glyceraldehyde phosphate dehydrogenase (GAPDH) were commercially synthesized using either previously published oligonucleotide sequence or ovine gene sequence (Sigma-Genosys). Forward and reverse primers were as follows: 5'-CAC CAA AAC CCT CAT CAA GAC 3'-and 5'-GAT GAA GTC CAA ACC AGT GAC-3', respectively, for leptin (GenBank accession no. AY01469351); 5'-AGC ATA CAG GTC CTG GCA TCT-3' and 5'-TGC CAT CCA ACC ACT CAG TA-3', respectively, for cyclophilin (GenBank accession no. AY251270); and 5'-GGC GTG AAC CAC GAG AAG TAT AA-3' and 5'-CCC TCC ACG ATG CCA AAG T-3', respectively, for GAPDH (housekeeping gene) (25). Maternal adipose cDNA dilutions were used to generate standard curves for GAPDH and leptin amplification. Real-time PCR was performed using SYBR Green JumpStart Taq ReadyMix (Sigma). The reaction consisted of 2 µl cDNA (fetal PAT at 1:200 dilution and placenta at 1:10 dilution), 12.5 µl SYBR Green Ready Mix, 0.2 µM of each primer, and diethylpyrocarbonate-H2O to a final volume of 25 µl. Reactions were performed in triplicate on a 96-well plate. The real-time PCR was performed using an ABI 7000 (Applied Biosystems, Warrington, UK). Samples were first heated to 95 C for 5 min, and the PCR cycle consisted of 95 C for 15 sec and 60 C for 1 min. This cycle was repeated 40 times for all cDNA amplification. Specificity of amplification was determined by melting curve analysis, and PCR efficiency for each set of primers was calculated using the following equation: efficiency (E) = 10(–1/slope) – 1. This was calculated to be 115% for leptin, 113% for GAPDH, and 134% for cyclophilin. Variation between assays was determined by analysis of a reference sample included in all runs; the interassay coefficient of variation was 9%. Data analysis was performed using Sequence Detection System software (SDS; Applied Biosystems). Levels of leptin mRNA were first normalized to GAPDH expression and then expressed as a fold change from the mean value of the respective control group measured in the same assay. No changes in the expression of GAPDH in PAT or cyclophilin in placenta were observed with gestational age or treatment.

Statistical analysis
All data are presented as mean ± SEM values. Data from the infused fetuses were compared by either one-way ANOVA or two-way ANOVA with repeated measures, followed by Tukey’s test as appropriate. Data from the dexamethasone-treated animals and from the AX and TX fetuses were compared with values from their respective control groups by unpaired t test. Differences at P < 0.05 were regarded as significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Localization of leptin protein and mRNA
Leptin protein and mRNA were localized in maternal PAT, fetal PAT, and placenta in control animals at 130 d of gestation. In maternal and fetal PAT, leptin protein was detected evenly across the sections and was localized to the periphery of the adipocytes (Fig. 1Go, B and C). Expression of leptin mRNA was identified in maternal and fetal PAT in a similar pattern to the leptin protein (Fig. 1Go, E and F).


Figure 1
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  		FIG. 1. Localization of leptin protein and mRNA in maternal PAT, fetal PAT, and placenta at 130 d of gestation. A, Negative control; B, C, and D, leptin protein localization in maternal PAT, fetal PAT, and placenta, respectively; E, F, and G, leptin mRNA localization in maternal PAT, fetal PAT, and placenta, respectively; H, I, and J, control hybridization with sense probes in maternal PAT, fetal PAT, and placenta, respectively. Scale bars, A and B, 100 µm; C–J, 50 µm.

 
In the ovine placenta at 130 d of gestation, leptin protein and mRNA were present on the membranes of the feto-maternal interdigitating villi (Fig. 1Go, D and G). However, it was not possible to distinguish whether staining was localized on the fetal trophoblast layer or the feto-maternal syncytiotrophoblast layer. The sense probes showed negligible staining for leptin mRNA in all tissues (Fig. 1Go, H–J). Similar localization of leptin protein and mRNA were observed in all three tissues taken from animals at 144 d of gestation.

Effect of gestational age
In the intact control fetuses, plasma concentrations of cortisol, T3, and leptin, but not T4, increased significantly between 130 and 140 d of gestation (P < 0.05) (Table 1Go and Fig. 2AGo). Over this period of gestation, the developmental change in plasma leptin concentration was associated with a significant rise in leptin mRNA abundance in fetal PAT. Leptin mRNA expression in fetal PAT at 144 d of gestation was significantly greater than at 130 d of gestation (P < 0.05) (Fig. 2BGo). Between 130 and 144 d of gestation, there were no significant differences in either absolute or relative PAT weights (Table 2Go). In the placenta, although leptin protein and mRNA expression were identified by immunohistochemistry and in situ hybridization at 130 and 144 d of gestation, leptin mRNA abundance was very low and did not change with gestational age (data not shown).


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  		TABLE 1. Mean ± SEM plasma concentrations of cortisol, T4, and T3 in the animals of the different treatment groups

 

Figure 2
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  		FIG. 2. Mean ± SEM values of plasma leptin (A) and adipose leptin mRNA (B) in intact fetuses at 130 d and intact, AX, and TX fetuses at 144 d of gestation. *, P < 0.05, significant difference from intact fetuses at 144 d.

 

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  		TABLE 2. Mean ± SEM values of body weight and absolute PAT and relative PAT weights, at delivery in the fetuses of the different treatment groups

 
Effect of manipulation of plasma glucocorticoid concentration
Fetal adrenalectomy, which prevented the prepartum surges in both plasma cortisol and T3, abolished the gestational rise in circulating leptin concentration (Table 1Go and Fig. 2AGo). At 144 d of gestation, AX fetuses had significantly lower plasma concentrations of cortisol, T3, and leptin compared with intact fetuses (P < 0.05) (Table 1Go and Fig. 2AGo). However, there was no significant difference in plasma T4 concentration between the two groups of fetuses (Table 1Go). The gestational rise in leptin mRNA abundance in PAT was also prevented in the AX fetuses; at 144 d of gestation, mean leptin mRNA abundance in the AX fetuses was significantly lower than in the intact fetuses (P < 0.05) (Fig. 2BGo). There were no significant differences in absolute or relative PAT weights between the intact and AX fetuses at 144 d of gestation (Table 2Go).

Exogenous infusion of cortisol into the fetus for 5 d caused significant increases in plasma concentrations of cortisol, T3, and leptin but not T4 (Table 1Go and Fig. 3AGo). Compared with the pretreatment baseline, a significant rise in plasma leptin concentration was observed on all days of the infusion in the cortisol-treated fetuses (P < 0.05) (Fig. 3AGo). Furthermore, within 24 h of the start of the infusion and on each day thereafter, plasma leptin in the cortisol-infused fetuses was significantly higher than in those infused with saline (P < 0.05) (Fig. 3AGo). The cortisol-induced rise in plasma leptin was associated with a significant increase in leptin mRNA expression in fetal PAT; on the fifth day of infusion, leptin mRNA abundance was significantly higher in the cortisol-infused fetuses compared with the saline-infused fetuses (P < 0.05) (Fig. 3BGo). Absolute and relative PAT weights were similar in the saline- and cortisol-infused fetuses (Table 2Go).


Figure 3
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  		FIG. 3. Mean ± SEM values of plasma leptin concentration (A) and adipose leptin mRNA (B) in the saline-, cortisol-, and T3-infused fetuses. For plasma leptin, within each day, columns with different letters are significantly different from each other (P < 0.05), and, within each treatment group, * indicates a significant difference from preinfusion baseline (P < 0.05). For adipose leptin mRNA on the fifth day of infusion, {dagger} indicates a significant difference from saline-infused fetuses (P < 0.05).

 
In the fetus, maternal dexamethasone treatment caused significant increments in both plasma leptin and leptin mRNA in PAT (P < 0.05) (Fig. 4AGo). These changes were associated with a reduction in plasma cortisol, an increase in plasma T3 and no change in plasma T4, or absolute and relative PAT weights, in fetuses of the dexamethasone-treated ewes compared with those whose mothers were treated with saline (Tables 1Go and 2Go). In the ewe, dexamethasone treatment had no effect on either plasma leptin or PAT leptin mRNA abundance but reduced circulating concentrations of cortisol, T3, and T4 (Table 1Go and Fig. 4BGo).


Figure 4
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  		FIG. 4. Mean ± SEM values of plasma leptin and adipose leptin mRNA in fetuses (A) and ewes (B) sampled at 10 h after the second daily injection of either saline or dexamethasone. *, P < 0.05, significant difference from the saline-treated animals.

 
Effect of manipulation of plasma thyroid hormone concentration
At 144 d of gestation, plasma leptin and PAT leptin mRNA levels were significantly higher in TX fetuses than in intact fetuses (P < 0.05) (Fig. 2Go, A and B). In the TX fetuses, plasma concentrations of T3 and T4 were significantly lower than in the intact fetuses, but plasma cortisol remained similar (P < 0.05) (Table 1Go). Fetal body weight was significantly lower and relative PAT weight was significantly higher in the TX fetuses compared with the intact fetuses at 144 d (P < 0.05) (Table 2Go).

An exogenous infusion of T3 in utero for 5 d had no effect on plasma leptin or leptin mRNA abundance in fetal PAT (Fig. 3Go). There was no significant difference in plasma leptin concentration between the T3 and saline-infused fetuses on any day of the infusion (Fig. 3AGo). Furthermore, on the fifth day of the infusion, PAT leptin mRNA abundance and absolute and relative PAT weights in the T3-infused fetuses were similar to those seen in fetuses treated with saline (Fig. 3BGo and Table 2Go).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The present study demonstrates that circulating leptin and PAT leptin mRNA levels are regulated in utero by glucocorticoids and thyroid hormones. The normal increments in plasma leptin and PAT leptin mRNA seen in the sheep fetus near term were abolished by fetal adrenalectomy, and both endogenous and synthetic glucocorticoids induced up-regulation of PAT leptin gene expression and, in turn, plasma leptin concentration. Therefore, the developmental rise in leptin production may be caused, at least in part, by the prepartum cortisol surge and can be induced prematurely by antenatal glucocorticoid treatment. In contrast, thyroid hormones had an inhibitory effect on PAT leptin synthesis and secretion in fetal sheep close to term. Although exogenous T3 infusion earlier in gestation had no effect on circulating leptin or PAT leptin mRNA, hypothyroidism in utero was associated with elevated circulating leptin and PAT leptin mRNA abundance. Therefore, in the ovine fetus during late gestation, ontogenic changes in PAT leptin production may result from the balance of stimulatory and inhibitory influences of the glucocorticoids and thyroid hormones, respectively.

Developmental and endocrine-induced changes in plasma leptin concentration observed in the fetuses of the present study were associated with changes in PAT leptin gene expression. This is in agreement with previous findings in normally fed and undernourished sheep in which circulating leptin in utero correlated with leptin mRNA abundance in fetal PAT (11). However, in the current study, the magnitude of change in PAT leptin mRNA abundance observed with gestational age and manipulation of glucocorticoid and thyroid hormone concentrations was disproportionately greater than the coincident change in plasma leptin concentration. These findings may be attributable to smaller subsequent effects on leptin protein synthesis and/or secretion from PAT, the relative clearance of leptin from the fetal circulation, or reductions in leptin synthesis by other fetal and placental tissues. In the sheep fetus near term, 85–95% of adipose tissue is perirenal (26), and leptin mRNA abundance is greater in PAT than in other adipose depots (5). Therefore, PAT appears to be an important source of circulating leptin in the ovine fetus during late gestation. Indeed, as demonstrated in the present and previous studies (5, 27), the ovine placenta does not express significant amounts of leptin mRNA and is unlikely to make a major contribution to leptin secretion in utero.

Before birth, glucocorticoids stimulated leptin gene expression in fetal PAT. These findings extend those that previously showed glucocorticoid-induced changes in plasma leptin in utero (13) but contrast with an earlier investigation, using tissues collected by our laboratory, that showed no change in PAT leptin mRNA abundance in sheep fetuses toward term or after glucocorticoid manipulation (28). Differences between these studies may be attributable to the more sensitive method of measurement of gene expression used in the current study. Glucocorticoids are likely to act directly on leptin gene transcription in utero because a glucocorticoid response element has been identified in the promoter region of the human leptin gene (29). In addition, the pronounced leptin response seen in the dexamethasone-exposed fetuses may have been partly caused by a coincident rise in plasma insulin concentration that is a known stimulant of leptin production in adipose tissue (30, 31). The actions of dexamethasone in utero are consistent with previous studies that show higher umbilical leptin concentration in preterm human infants whose mothers were treated with antenatal glucocorticoids (32). In the pregnant ewe, however, the lack of response to dexamethasone contrasts with previous observations in adult nonpregnant rats and human subjects in which dexamethasone administration, at higher and lower doses to the present study, increased adipose leptin mRNA and plasma leptin levels (33, 34).

The stimulatory effects of glucocorticoids on plasma leptin and PAT leptin mRNA levels in utero did not appear to be mediated by changes in thyroid hormone concentration, unlike that seen in other physiological systems in the fetus (14, 15). In contrast, the thyroid hormones had an inhibitory effect on PAT leptin mRNA abundance and plasma leptin concentration in the sheep fetus near term. Fetal thyroidectomy, which lowered plasma T4 concentration to an undetectable level and prevented the prepartum T3 surge, caused elevations in both PAT leptin gene expression and circulating leptin concentration. Previous studies have demonstrated an inverse relationship between plasma concentrations of leptin and T3 in rats after experimental manipulation of thyroid hormone status and in human subjects with thyroid dysfunction (35, 36, 37).

There are a number of mechanisms by which thyroid hormones in utero may influence leptin synthesis and secretion by fetal adipose tissue. To date, a thyroid hormone response element has not been identified in the leptin gene, although T3 causes dose-dependent inhibition of leptin gene expression and protein secretion in brown and white adipocytes cultured from adult rats (38). There may also be a direct action of the high TSH concentration associated with thyroidectomy, because TSH increases leptin secretion from human adipose tissue in vitro (39). In addition, some of the effects of the thyroid hormones on leptin synthesis and secretion in utero may be induced indirectly by changes in tissue sensitivity to sympathetic stimulation. In adult animals, ß3-adrenergic stimulation inhibits leptin gene expression in adipose tissue and suppresses circulating leptin concentration (40, 41). Furthermore, adipose ß3-adrenergic receptor mRNA abundance has been shown to depend on thyroid hormones in adult rats (42). In sheep fetuses, noradrenergic-induced cellular respiration is lower in PAT obtained from TX fetuses compared with TX fetuses infused with a replacement dose of T3 (43), which suggests that thyroid hormones have an important role in determining adrenergic sensitivity in fetal adipose tissue. Hypothyroidism in the ovine fetus is also known to alter oxygen consumption (23), and this, in turn, may have had consequences for the control of leptin synthesis in PAT. Although arterial blood oxygen tension remains unchanged in TX fetuses (23), chronic hypoxia in utero increases both leptin gene expression in periadrenal fat and circulating leptin concentration (44). Last, part of the rise in circulating leptin concentration seen in the TX fetuses may have been attributable to greater relative PAT weight. The TX fetuses were growth retarded compared with intact fetuses at the same gestational age, but absolute PAT weight was maintained. Previous studies in fetal sheep have shown that plasma leptin concentration correlates with PAT mass and specifically with the relative amount and cell size of unilocular fat (26). However, the cellular structure of PAT was not investigated in the fetuses of the present study. Therefore, there may be both direct and indirect effects of thyroid hormones on leptin synthesis in utero. In the present study, although thyroid hormone deficiency upregulated leptin synthesis in the sheep fetus near term, an exogenous infusion of T3 between 125 and 130 d of gestation had no effect on PAT leptin mRNA abundance or plasma leptin concentration. This may have been attributable to a relative lack of thyroid hormone receptors and/or postreceptor sensitivity in fetal PAT at this gestational age. Furthermore, indirect mechanisms of thyroid hormone action on leptin synthesis may be influenced by gestational age and/or the experimental method used to manipulate thyroid hormone concentration in utero. These include PAT sensitivity to catecholamines and the control of TSH, sympathetic output, oxygen consumption, and relative PAT mass.

Developmental and glucocorticoid-induced increments in plasma leptin concentration may have an important role in the control of fetal maturation near term. Indeed, some of the maturational effects of endogenous and synthetic glucocorticoids in the ovine fetus may be mediated by the coincident rise in circulating leptin. Leptin receptors have been identified in a variety of fetal and placental tissues in several animal species (6, 7). For example, leptin receptor mRNA and protein are expressed in the lungs of baboon fetuses (45), and leptin has been shown to stimulate surfactant production in type II pneumocytes cultured from fetal rats and rabbits (46, 47). At birth, thyroid hormone and catecholamine release in response to labor and delivery into a cold environment may contribute to the fall in plasma leptin seen in human and ovine neonates (13, 48). This may be an important drive to suckling in the immediate postnatal period. Furthermore, the role of perinatal and endocrine-induced changes in leptin production in the long-term control of food intake and energy balance remains to be established. Exposure of the fetus to leptin at critical periods of development may have important consequences for the formation and activity of hypothalamic neural networks responsible for appetite regulation and, hence, obesity in adult life (1, 2).


   Acknowledgments
 
We are grateful to Margaret Blackberry (University of Western Australia) for the measurement of plasma leptin and to the members of the Department of Physiology, Development, and Neuroscience who provided technical assistance in this study.


   Footnotes
 
The study was supported by the Biotechnology and Biological Sciences Research Council.

Disclosure Statement: The authors have nothing to disclose.

First Published Online May 10, 2007

Abbreviations: AX, Adrenalectomized; DIG, digoxigenin; GAPDH, glyceraldehyde phosphate dehydrogenase; PAT, perirenal adipose tissue; SSC, sodium citrate; TBS, Tris-buffered saline; TX, thyroidectomized.

Received March 7, 2007.

Accepted for publication May 1, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. McMillen IC, Adam CL, Muhlhausler BS 2005 Early origins of obesity: programming the appetite regulatory system. J Physiol 565:9–17[Abstract/Free Full Text]
  2. Bouret SG, Simerly RB 2006 Developmental programming of hypothalamic feeding circuits. Clin Genet 70:295–301[CrossRef][Medline]
  3. Bouret SG, Draper SJ, Simerly RB 2004 Trophic action of leptin on hypothalamic neurons that regulate feeding. Science 304:108–110[Abstract/Free Full Text]
  4. Jaquet D, Leger J, Levy-Marchal C, Oury JF, Czernichow P 1998 Ontogeny of leptin in human fetuses and newborns: effect of intrauterine growth retardation on serum leptin concentrations. J Clin Endocrinol Metab 83:1243–1246[Abstract/Free Full Text]
  5. Ehrhardt RA, Bell AW, Boisclair YR 2002 Spatial and developmental regulation of leptin in fetal sheep. Am J Physiol 282:R1628–R1635
  6. Hoggard N, Hunter L, Duncan JS, Williams LM, Trayhurn P, Mercer JG 1997 Leptin and leptin receptor mRNA and protein expression in the murine fetus and placenta. Proc Natl Acad Sci USA 94:11073–11078[Abstract/Free Full Text]
  7. Buchbinder A, Lang U, Baker RS, Khoury JC, Mershon J, Clark KE 2001 Leptin in the ovine fetus correlated with fetal and placental size. Am J Obstet Gynecol 185:786–791[CrossRef][Medline]
  8. Helland IB, Reseland JE, Saugstad OD, Drevon CA 1998 Leptin levels in pregnant women and newborn infants: gender differences and reduction during the neonatal period. Pediatrics 101:E12
  9. Varvarigou A, Mantzoros CS, Beratis NG 1999 Cord blood leptin concentrations in relation to intrauterine growth. Clin Endocrinol 50:177–183[CrossRef][Medline]
  10. Yuen BSJ, McMillen IC, Symonds ME, Owens PC 1999 Abundance of leptin mRNA in fetal adipose tissue is related to fetal body weight. J Endocrinol 163:R1–R4
  11. Yuen BSJ, Owens PC, McFarlane JR, Symonds ME, Edwards LJ, Kauter KG, McMillen IC 2002 Circulating leptin concentrations are positively related to leptin messenger RNA expression in the adipose tissue of fetal sheep in the pregnant ewe fed at or below maintenance energy requirements during late gestation. Biol Reprod 67:911–916[Abstract/Free Full Text]
  12. Ashworth CJ, Hoggard N, Thomas L, Mercer JG, Wallace JM, Lea RG 2000 Placental leptin. Rev Reprod 5:18–24[Abstract]
  13. Forhead AJ, Thomas L, Crabtree J, Hoggard N, Gardner DS, Giussani DA, Fowden AL 2002 Plasma leptin concentration in fetal sheep during late gestation: ontogeny and effect of glucocorticoids. Endocrinology 143:1166–1173[Abstract/Free Full Text]
  14. Forhead AJ, Fowden AL 2002 Effects of thyroid hormones on pulmonary and renal angiotensin-converting enzyme concentrations in fetal sheep near term. J Endocrinol 173:143–150[Abstract]
  15. Forhead AJ, Poore KR, Mapstone J, Fowden AL 2003 Developmental regulation of hepatic and renal gluconeogenic enzymes by thyroid hormones in fetal sheep during late gestation. J Physiol 548:941–947[Abstract/Free Full Text]
  16. Forhead AJ, Curtis K, Kaptein E, Visser TJ, Fowden AL 2006 Developmental control of iodothyronine deiodinases by cortisol in the ovine fetus and placenta. Endocrinology 147:5988–5994[Abstract/Free Full Text]
  17. Barnes RJ, Comline RS, Silver M 1977 The effects of bilateral adrenalectomy or hypophysectomy of the foetal lamb in utero. J Physiol 264:429–447[Abstract/Free Full Text]
  18. Hopkins PS, Thorburn GD 1972 The effects of foetal thyroidectomy on the development of the ovine foetus. J Endocrinol 54:55–66[Medline]
  19. Comline RS, Silver M 1972 The composition of foetal and maternal blood during parturition in the ewe. J Physiol 222:248–256
  20. Forhead AJ, Li J, Saunders JC, Dauncey MJ, Gilmour RS, Fowden AL 2000 Control of ovine hepatic growth hormone receptor and insulin-like growth factor-I by thyroid hormones in utero. Am J Physiol 278:E1166–E1174
  21. Blache D, Tellam RL, Chagas LM, Blackberry MA, Vercoe PE, Martin GB 2000 Level of nutrition affects leptin concentrations in plasma and cerebrospinal fluid in sheep. J Endocrinol 165:625–637[Abstract]
  22. Robinson PM, Comline RS, Fowden AL, Silver M 1983 Adrenal cortex of fetal lamb: changes after hypophysectomy and effects of Synacthen on cytoarchitecture and secretory activity. Q J Exp Physiol 68:15–27[Abstract/Free Full Text]
  23. Fowden AL, Silver M 1995 The effects of thyroid hormones on oxygen and glucose metabolism in the sheep fetus during late gestation. J Physiol 482:203–213
  24. Chomczynski P, Sacchi N 1987 Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem 162:156–159[Medline]
  25. Budhia S, Haring LF, McConnell I, Blacklaws BA 2006 Quantitation of ovine cytokine mRNA by real-time RT-PCR. J Immunol Methods 309:160–172[CrossRef][Medline]
  26. Muhlhausler BS, Roberts CT, Yuen BSJ, Marrocco E, Budge H, Symonds ME, McFarlane JR, Kauter KG, Stagg P, Pearse JK, McMillen IC 2003 Determinants of fetal leptin synthesis, fat mass, and circulating leptin concentrations in well-nourished ewes in late pregnancy. Endocrinology 144:4947–4954[Abstract/Free Full Text]
  27. Thomas L, Wallace JM, Aitken RP, Mercer JG, Trayhurn P, Hoggard N 2001 Circulating leptin during ovine pregnancy in relation to maternal nutrition, body composition and pregnancy outcome. J Endocrinol 169:465–476[Abstract]
  28. Mostyn A, Pearce S, Budge H, Elmes M, Forhead AJ, Fowden AL, Stephenson T, Symonds ME 2003 Influence of cortisol on adipose tissue development in the fetal sheep during late gestation. J Endocrinol 176:23–30[Abstract]
  29. Gong D-W, Bi S, Pratley RE, Weintraub BD 1996 Genomic structure and promoter analysis of the human obese gene. J Biol Chem 271:3971–3974[Abstract/Free Full Text]
  30. Wabitsch M, Jensen PB, Blum WF, Christoffersen CT, Englaro P, Heinze E, Rascher W, Teller W, Tornqvist H, Hauner H 1996 Insulin and cortisol promote leptin production in cultured human fat cells. Diabetes 45:1435–1438[Abstract]
  31. Franko KL, Giussani DA, Forhead AJ, Fowden AL 2007 Effects of dexamethasone on the glucogenic capacity of fetal, pregnant and non-pregnant adult sheep. J Endocrinol 192:67–73[Abstract/Free Full Text]
  32. Shekhawat PS, Garland JS, Shivpuri C, Mick GJ, Sasidharan P, Pelz CJ, McCormick KL 1998 Neonatal cord blood leptin: its relationship to birth weight, body mass index, maternal diabetes, and steroids. Pediatr Res 43:338–343[Medline]
  33. De Vos P, Saladin R, Auwerx J, Staels B 1995 Induction of ob gene expression by corticosteroids is accompanied by body weight loss and reduced food intake. J Biol Chem 270:15958–15961[Abstract/Free Full Text]
  34. Papaspyrou-Rao S, Schneider S, Petersen RN, Fried SK 1997 Dexamethasone increases leptin expression in humans in vivo. J Clin Endocrinol Metab 82:1635–1637[Abstract/Free Full Text]
  35. Leonhardt U, Gerdes E, Ritzel U, Schafer G, Becker W, Ramadori G 1999 Immunoreactive leptin and leptin mRNA expression are increased in rat hypo- but not hyperthyroidism. J Endocrinol 163:115–121[Abstract]
  36. Zimmermann-Belsing T, Brabant G, Holst JJ, Feldt-Rasmussen U 2003 Circulating leptin and thyroid dysfunction. Eur J Endocrinol 149:257–271[Abstract]
  37. Zabrocka L, Klimek J, Swierczynski J 2006 Evidence that triiodothyronine decreases rat serum leptin concentration by down-regulation of leptin gene expression in white adipose tissue. Life Sci 79:1114–1120[CrossRef][Medline]
  38. Medina-Gomez G, Calvo R-M, Obregon M-J 2004 T3 and Triac inhibit leptin secretion and expression in brown and white rat adipocytes. Biochim Biophys Acta 1682:38–47[Medline]
  39. Menendez C, Baldelli R, Camina JP, Escudero B, Peino R, Dieguez C, Casanueva FF 2003 TSH stimulates leptin secretion by a direct effect on adipocytes. J Endocrinol 176:7–12[Abstract]
  40. Moinat M, Deng C, Muzzin P, Assimacopoulos-Jeannet F, Seydoux J, Dulloo AG, Giacobino J-P 1995 Modulation of obese gene expression in rat brown and white adipose tissue. FEBS Lett 373:131–134[CrossRef][Medline]
  41. Trayhurn P, Duncan JS, Rayner DV, Hardie LJ 1996 Rapid inhibition of ob gene expression and circulating leptin levels in lean mice by the ß3-adrenoceptor agonists BRL 35135A and ZD2079. Biochem Biophys Res Commun 228:605–610[CrossRef][Medline]
  42. Fain JN, Coronel EC, Beauchamp MJ, Bahouth SW 1997 Expression of leptin and ß3-adrenergic receptors in rat adipose tissue in altered thyroid states. Biochem J 322:145–150[Medline]
  43. Klein AH, Reviczky A, Padbury JF 1984 Thyroid hormones augment catecholamine-stimulated brown adipose tissue thermogenesis in the ovine fetus. Endocrinology 114:1065–1069[Abstract]
  44. Ducsay CA, Hyatt K, Mlynarczyk M, Kaushal KM, Myers DA 2006 Long-term hypoxia increases leptin receptors and plasma leptin concentrations in the late-gestation ovine fetus. Am J Physiol 291:R1406–R1413
  45. Hensen MC, Swan KF, Edwards DE, Hoyle GW, Purcell J, Castracane VD 2004 Leptin receptor expression in fetal lung increases in late gestation in the baboon: a model for human pregnancy. Reproduction 127:87–94[Abstract/Free Full Text]
  46. Bergen HT, Cherlet TC, Manuel P, Scott JE 2002 Identification of leptin receptors in lung and isolated fetal type II cells. Am J Respir Cell Mol Biol 27:71–77[Abstract/Free Full Text]
  47. Torday JS, Sun H, Wang L, Torres E, Sunday ME, Rubin LP 2002 Leptin mediates the parathyroid hormone-related protein paracrine stimulation of fetal lung maturation. Am J Physiol 282:L405–L410
  48. Schubring C, Siebler T, Englaro P, Blum WF, Kratzsch J, Triep K, Kiess W 1998 Rapid decline of serum leptin levels in healthy neonates after birth. Eur J Pediatr 157:265–271[CrossRef][Medline]



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